ORIGINAL ARTICLE

Latent and active aurone synthase from petals of C. grandiflora:a polyphenol oxidase with unique characteristics

Christian Molitor • Stephan Gerhard Mauracher •

Sanela Pargan • Rupert L. Mayer • Heidi Halbwirth •

Annette Rompel

Received: 30 October 2014 /Accepted: 3 February 2015 / Published online: 20 February 2015

� The Author(s) 2015. This article is published with open access at Springerlink.com

Abstract

Main conclusion Aurone synthase belongs to the novel

group 2 polyphenol oxidases and the presented kinetic

characterization suggests a differing aurone biosynthe-

sis in Asteraceae species compared to snapdragon.

Aurone synthases (AUS) are polyphenol oxidases (PPO)

physiologically involved in the formation of yellow aurone

pigments in petals of various Asteraceae species. They

catalyze the oxidative conversion of chalcones into au-

rones. Latent (58.9 kDa) and active (41.6 kDa) aurone

synthase from petals of C. grandiflora was purified by a

quantitative removal of pigments using aqueous two-phase

separation and several subsequent chromatographic steps.

The purified enzymes were identified as cgAUS1

(A0A075DN54) and sequence analysis revealed that

cgAUS1 is a member of a new group of plant PPOs. Mass

determination experiments of intact cgAUS1 gave evi-

dence that the C-terminal domain, usually shielding the

active site of latent polyphenol oxidases, is linked to the

main core by a disulfide bond. This is a novel and unique

structural feature of plant PPOs. Proteolytic activation

in vivo leads to active aurone synthase possessing a resi-

dual peptide of the C-terminal domain. Kinetic charac-

terization of purified cgAUS1 strongly suggests a specific

involvement in 4-deoxyaurone biosynthesis in Coreopsis

grandiflora (Asteraceae) that differs in various aspects

compared to the 4-hydroxyaurone formation in Antir-

rhinum majus (Plantaginaceae): cgAUS1 is predicted to be

localized in the thylakoid lumen, it possesses exclusively

diphenolase activity and the results suggest that aurone

formation occurs at the level of chalcone aglycones. The

latent enzyme exhibits allosteric activation which changes

at a specific product concentration to a constant reaction

rate. The presented novel structural and functional prop-

erties of aurone synthase provide further insights in the

diversity and role of plant PPOs.

Keywords Allosteric activation � Aurone synthase �4-Deoxyaurone � Polyphenol oxidase � Proteolyticactivation � Type-3 copper enzyme

Abbreviations

AEX Anion exchange

ATPS Aqueous two-phase separation

cgAUS Coreopsis grandiflora aurone synthase

CEX Cation exchange

CV Column volumes

DTT DL-Dithiothreitol

MWCO Molecular weight cut off

PPO(s) Polyphenol oxidase(s)

vvCO Vitis vinifera catechol oxidase

Special topic:Polyphenols: biosynthesis and function in plants and

ecosystems.Guest editor: Stefan Martens.

Electronic supplementary material The online version of thisarticle (doi:10.1007/s00425-015-2261-0) contains supplementarymaterial, which is available to authorized users.

C. Molitor � S. G. Mauracher � S. Pargan � A. Rompel (&)

Institut fur Biophysikalische Chemie, Fakultat fur Chemie,

Universitat Wien, Althanstraße 14, 1090 Vienna, Austria

e-mail: annette.rompel@univie.ac.at

R. L. Mayer

Department of Analytical Chemistry, University of Vienna,

Wahringer Straße 38, 1090 Vienna, Austria

H. Halbwirth

Institute of Chemical Engineering, University of Technology

Vienna, Getreidemarkt 9, 1060 Vienna, Austria

123

Planta (2015) 242:519–537

DOI 10.1007/s00425-015-2261-0

Introduction

Chalcones and aurones are yellow plant pigments related to

flavonoids. They are called anthochlor pigments and are

found in yellow flowers of Asteraceae species, snapdragon

(Antirrhinum majus L.) and carnations (Dianthus

caryophyllus) (Harborne 1967). Two types of anthochlors

exist distinguished by the presence or absence of a hydroxyl

group in ring A at position 60 of chalcones and position 4 ofaurones (note differing numbering of chalcones and au-

rones, Fig. 1). In Asteraceae, anthochlor pigments of the

deoxy type are typically found. Several 60-deoxychalconesand 4-deoxyaurones (Fig. 1) were identified in extracts of

petals of the genus of Coreopsis (C. maritima, C. gigantea,

C. bigelovii, C. grandiflora, C. mutica, C. tinctoria, C.

saxicola and C. lanceolata) (Geissman and Heaton 1943,

1944; Geissman and Moje 1951; Shimokoriyama and Hat-

tori 1953; Geissman et al. 1956; Harborne and Geissman

1956; Crawford 1970; Nicholls and Bohm 1979; Crawford

and Smith 1980, 1983; Tanimoto et al. 2009; Shang et al.

2013; Okada et al. 2014; Wang et al. 2014). In contrast to

the extensively studied flavonoid formation, knowledge

about aurone biosynthesis is still very limited.

Aurone formation was studied primarily in Antirrhinum

majus (snapdragon) as a model plant for the formation of

hydroxyanthochlors. A homologue of polyphenol oxidases

(PPOs), aureusidin synthase from A. majus (AmAS1), was

identified as the key enzyme in aurone biosynthesis in

snapdragon (Nakayama et al. 2000) as one of the rare ex-

amples of a PPO involved in an anabolic pathway (Strack

and Schliemann 2001).

PPOs are type-3 copper enzymes which are widespread

among plants, fungi and bacteria displaying a broad range of

putative physiological roles, primarily in the protection of

organisms against biotic and abiotic stress. A specific in-

volvement in biosynthetic processes, beside aurone biosyn-

thesis, was shown only in a few cases (Mayer 2006; Wahler

et al. 2009;Araji et al. 2014). Plant PPOs are expressed in their

latent state, in which the active site of the enzyme is shielded

by the C-terminal domain, which is ultimately removed by

proteolytic cleavage resulting in a fully active enzyme (King

and Flurkey 1987; Espin et al. 1999; Gandia-Herrero et al.

2005; Marusek et al. 2006; Flurkey and Inlow 2008). It was

shown that the latent pro-enzymes undergo allosteric activa-

tion with several substrates (Valero and Garcia-Carmona

1992, 1998; Nillius et al. 2008). In vitro, the pro-enzymes can

be activated by proteases, an acidic pH, fatty acids or deter-

gents (e.g. SDS) (Yoruk and Marshall 2003).

PPOs comprise monophenolases [tyrosinases (EC

1.14.18.1)], catalyzing the o-hydroxylation of monophe-

nols and oxidation of o-diphenols to o-quinones, and

diphenolases [catechol oxidases (EC 1.10.3.1)], which lack

monophenolase activity and catalyze the oxidation reaction

exclusively. Although PPOs have been studied over many

decades and crystal structures of tyrosinases and catechol

oxidases are available, no prediction of their reactivity

based on sequence information has been possible so far.

During catalysis, PPOs can undergo suicide inactivation,

which is proposed to be caused by a substrate docking

mode that leads to the reductive loss of copper (Land et al.

2008; Munoz-Munoz et al. 2008, 2011, 2012a, b; Ramsden

et al. 2009; Ramsden and Riley 2010a).

Fig. 1 60-deoxychalcones and 4-deoxyaurones identified in petals of

C. maritima, C. gigantea, C. bigelovii, C. grandiflora, C. mutica, C.

tinctoria, C. saxicola and C. lanceolata (Geissman and Heaton 1943,

1944; Geissman and Moje 1951; Shimokoriyama and Hattori 1953;

Geissman et al. 1956; Harborne and Geissman 1956; Crawford 1970;

Nicholls and Bohm 1979; Crawford and Smith 1980, 1983; Tanimoto

et al. 2009; Shang et al. 2013; Okada et al. 2014; Wang et al. 2014)

520 Planta (2015) 242:519–537

123

Most plant PPOs contain an N-terminal chloroplast

transit peptide (cTP) and a thylakoid transfer domain and

are hence predicted to be transported to the thylakoid lu-

men in the chloroplast. Only a few PPOs lack the cTP, but

instead contain a N-terminal secretion signal peptide (Tran

et al. 2012). Apart from PPO from Populus trichocarpa,

AmAS1 is the only PPO for which a vacuolar localisation

was demonstrated (Ono et al. 2006b; Tran and Constabel

2011). In accordance with its presence in vacuoles, AmAS1

was found to be a glycoprotein (Nakayama et al. 2000).

Notably, a chalcone 20-O-glycosyltransferase is required

for the accumulation of aurones in transgenic plants

(Nakayama 2002; Shakya et al. 2012) because transport

into vacuoles occurs only on extensively modified struc-

tures and not on aglycones (Matern et al. 1986).

Recently, it was suggested that the formation of

4-deoxyaurones differs from that of 4-hydroxyaurones in

various aspects. Most notably, a PPO homologue lacking

the monophenolase activity appears to be responsible for

aurone formation (Miosic et al. 2013; Kaintz et al. 2014),

while the introduction of a second vicinal hydroxyl group

in the B-ring of chalcones is catalyzed in an independent

step by a chalcone 3-hydroxylase (Schlangen et al. 2010).

Recently, two cDNA clones (cgAUS1 (KC972611) and

cgAUS2 (KC878307)) of putative aurone synthases from

petals of Coreopsis grandiflora (cgAUS) were isolated

(Kaintz et al. 2014). cgAUS1 and cgAUS2 showed diver-

gent gene expression profiles. cgAUS1 was particularly

expressed in petals, as expected for a gene involved in the

biosynthesis of flower pigments. In contrast, cgAUS2 dis-

played higher expression rates in green leaves rather than

in flowers. Both gene products are predicted to be subcel-

lularly localized in the chloroplasts and possess highly

conserved motifs of plant PPOs (Kaintz et al. 2014).

To verify the putative involvement of cgAUS1 in aurone

biosynthesis in C. grandiflora, we purified high amounts of

active as well as latent cgAUS from petals for characterization

and subsequent crystallization experiments. The purified PPO,

identified as cgAUS1 (A0A075DN54), is a member of the

novel group 2 PPOs and possesses hitherto unknown structural

and functional properties. The substrate specificity of cgAUS1

is compared with a common catechol oxidase from Vitis

vinifera (vvCO). Based on our findings, we propose an aurone

biosynthetic pathway in Asteraceae species alternative to the

one reported forA.majus (Davies et al. 2006;Onoet al. 2006a).

Materials and methods

Plant material

Seedlings of C. grandiflora cv. Early sunrise were pur-

chased from Volmary (Munster, Germany), cultivated in

April 2011 in the experimental field Augarten of the

University of Vienna (Austria). Petals of all developmental

stages were harvested from June to September, all green

tissues were discarded and the yellow plant material was

shock-frozen in liquid nitrogen and stored at -80 �C.

Reagents

Butein and marein were purchased from Extrasynthesis

(Genay, France). All other used chemicals were purchased

from Sigma-Aldrich at the highest purity available.

Purification, yield and SDS-PAGE of catechol oxidase

from Vitis vinifera (vvCO)

Catechol oxidase from V. vinifera was used as a reference

enzyme to evaluate substrate specificity of aurone synthase.

The description of the isolation, purification, yield and

SDS-PAGE analysis of this enzyme is presented in Online

Resource Fig. S1.

Protein extraction and removal of pigments

The pigment removal by an aqueous two-phase separation

(ATPS) procedure described below is based on the methods

described by Sojo et al. (1998), which were, however,

significantly modified and optimized in several aspects.

Similar procedures were also reported for the isolation of

tyrosinase from Juglans regia (Zekiri et al. 2014) and

Agaricus bisporus (Mauracher et al. 2014). Protein ex-

traction was carried out on 6 kg petal tissue in total which

was processed in three batches as outlined below.

2 kg of frozen petals were homogenized with a hand-

held blender. 4 l buffer containing 125 mM sodium citrate,

pH 5.4, 4 % (v/v) Triton X-114, 0.5 % (w/v) sodium

ascorbate, 50 mM L-proline, 1 mM phenylmethylsulfonyl

fluoride (PMSF) and 2 mM benzamidine hydrochloride

were added and the pulp was centrifuged at 16,000g for

20 min at 4 �C. The supernatant was filtered through

cheesecloth, 15 g/l ammonium sulfate was added and the

solution was warmed up to 8–12 �C. This caused the so-

lution to become turbid due to micelle growth and subse-

quent liquid-liquid phase separation. After centrifugation at

16,000g for 10 min at 12 �C, the brownish detergent-rich

bottom phase was discarded and the brighter detergent-

poor top phase was supplemented with Triton X-114 to a

final concentration of approx. 4 % (v/v). The turbid solu-

tion was stirred for 15 min at 8–12 �C, and was then

centrifuged at 16,000g for 10 min at 12 �C. The bottom

phase was again discarded and ammonium sulfate was

added to a final concentration of 30 % saturation to the

orange top phase. After stirring to dissolve the salt, the

suspension was allowed to chill at 4 �C for 45 min. All the

Planta (2015) 242:519–537 521

123

following steps were performed at 4 �C. After centrifuga-tion at 16,000g for 45 min and filtration, PEG-4000 was

added to a final concentration of 4.5 % (w/v). The turbid

solution was centrifuged at 16,000g for 10 min and the

yellow–orange colored detergent-rich top phase was dis-

carded, while the lower detergent-poor phase was subjected

to another round of ATPS. This procedure was repeated

3–5 times with descending additions of PEG-4000 (4, 3.5,

3 % (w/v), respectively), until the red to yellow colored

pigments were removed quantitatively. To the resultant

clear and slightly beige colored enzyme solution ammo-

nium sulfate was added to a final concentration of approx.

85 % saturation and stored at 4 �C over night. The am-

monium sulfate solution beneath the floating pellet was

discarded with the pellet transferred to a bottle-top filter

(GE Healthcare, PES, 45 lm) to reduce the ammonium

sulfate solution content to a minimum via gravity-driven

filtration.

Enzymatic assay

AUS assays of initial purifications were performed by

monitoring the increase in absorbance at 415 nm due to the

conversion of butein into sulfuretin. The reaction mixture

contained 40 lM butein in 125 mM sodium citrate at pH 5.5.

Enzymatic assays for subsequent purifications were

performed by monitoring the oxidation of 50 lM fisetin in

125 mM sodium citrate at pH 5.5 at 280 nm (Jimenez et al.

1998). When allosteric activation during the enzymatic

reaction was visible, the fractions contained portions of

latent protein. Further assays for the latent forms were

performed either by adding 2.5 mM SDS to the assay

mixture or using 125 mM sodium citrate at pH 3.5 to ac-

tivate the protein to obtain a linear absorbance–time

relation.

Kinetic analysis

Experiments for determination of kinetic parameters of

aurone synthase were performed using sample 1 (cgAUS1)

while varying the substrate concentration. For evaluation of

the substrate specificity of aurone synthase, kinetic pa-

rameters of a purified catechol oxidase from Vitis vinifera

(vvCO) were determined as well. Mean values and standard

deviations were calculated based on four repetitions. Data

analysis was carried out by nonlinear regression (Mar-

quardt 1963). In cases where an accurate estimation of Km

by nonlinear fitting was not possible due to the low solu-

bility of substrates, the ratio kcat/Km was determined by

linear fitting to pseudo first-order kinetics ([S] � Km).

Separations of chalcones and aurones applying HPLC were

performed as previously described by Miosic et al. (2013)

using the protocol of Chandra et al. (2001).

Purified heterologously expressed cgAUS1 (Kaintz et al.

2014) exhibited properties identical to the latent cgAUS1

purified from the natural source. Due to traces of active

cgAUS1 in the sample purified from the natural source,

recombinantly expressed cgAUS1 was used to describe the

characteristics of the latent enzyme.

Protein purification by Fast Protein Liquid

Chromatography (FPLC)

All chromatographic purification steps were carried out

using an Akta Purifier (GE Healthcare) placed in a re-

frigerator to maintain 4 �C.An appropriate portion of the drained protein pellet

obtained was dissolved in 300 ml 20 mM sodium acetate

buffer (pH 5.0), centrifuged at 28,000g for 45 min at 4 �Cand further diluted with buffer until the conductivity of the

enzyme solution was decreased below 11 mS/cm. The so-

lution was applied to a SP-Sepharose FF column (HiScale

26/40, 70 ml bed volume) via a sample pump at a flow rate

of 13 ml/min. Proteins were eluted by an increasing sodi-

um chloride gradient over 15 column volumes (CV) up to

0.8 M sodium chloride at a flow rate of 5 ml/min. Although

the majority of the resulting fractions contained AUS ac-

tivity, slight maxima in activity could be determined. This

was taken into account for pooling of the active fractions.

The pooled fractions were diluted by a factor of three

with 20 mM sodium acetate (pH 5.0) and applied to a SP-

Sepharose HP column (XK 16, 10 ml bed volume) at a

flow rate of 5 ml/min. Elution was achieved by an in-

creasing sodium chloride gradient over 10 CV, up to

0.8 M at a flow rate of 3 ml/min. Active fractions were

pooled, concentrated by ultracentrifugation (30 kDa

molecular weight cut off, MWCO), diluted in 45 ml

30 mM Tris/HCl buffer (pH 8.5) and loaded on a Mono Q

HR 5/50 GL column. Proteins were eluted by an increasing

sodium chloride gradient up to 250 mM sodium chloride

over 200 CV. Several related fractions, showing either

ordinary activity or allosteric behavior towards fisetin as

substrate, were pooled, concentrated by ultracentrifugation

(30 kDa MWCO), diluted in 45 ml 20 mM sodium acetate

(pH 5.0) and loaded onto a Mono S HR 5/50 GL column.

Elution was performed by applying an increasing sodium

chloride gradient up to 500 mM over 200 CV. Selected

active and latent fractions were again subjected to Mono S

cation exchange chromatography at pH 5.0, whereas, due

to overlapping protein peaks, a change to 20 mM sodium

citrate (pH 5.6) was necessary as a polishing step for one

latent form.

A buffer exchange to 10 mM sodium acetate (pH 5.0) of

the purified fractions was performed by ultracentrifugation

(10 kDa MWCO) and the concentrated samples (6–10 mg/

ml) were stored at 4 �C.

522 Planta (2015) 242:519–537

123

Molecular mass determination

SDS-PAGE was performed according to the method of

Laemmli (1970), using Precision Plus Protein Standard

Dual Color (Bio-Rad) as the molecular weight marker. The

samples were applied to 10 % polyacrylamide gels under

reducing and non-reducing conditions by adding or ex-

cluding DL-dithiothreitol (DTT). Gels were stained with

Coomassie Brilliant Blue R-250.

Electrospray Ionization Mass Spectrometry (ESI–MS)

was performed on a nanoESI-QTOF mass spectrometer

(maXis 4G UHR-TOF, Bruker) with a mass resolving

power of about 40,000 in the used m/z—range and a mass

accuracy of better than 5 ppm (confirmed by standard

proteins) using a protein solution of 20 ll with a concen-

tration of approximately 1 lg/ll. Buffer exchange to

10 mM ammonium acetate (pH 5.0) of non-reduced sam-

ples was performed by ultracentrifugation. After buffer

exchange to 20 mM ammonium acetate (pH 7.5) by ultra-

centrifugation of samples for measurements under reducing

conditions, DTT was added to a final concentration of

50 mM. Subsequently, the samples were incubated for

45 min at 50 �C. Just prior to the measurements, acetoni-

trile to a final concentration of 25 % (v/v) and formic acid

to a final concentration of 0.05 % (v/v) were added.

Protein identification

Tryptic digestions of enzyme solutions containing ap-

proximately 10 lg protein were performed according to the

manufacturers’ description. The peptide samples were

dried by vacuum centrifugation and stored at -20 �C for

LC–MS/MS analysis. The samples were solubilized in 5 ll30 % (v/v) formic acid and diluted with 40 ll 2 % (v/v)

acetonitrile, 0.1 % (v/v) formic acid.

Analysis of the samples was carried out by

nanoUHPLC–ESI–MS/MS using a high-resolution orbitrap

mass spectrometer (Dionex Ultimate 3000 RSLCnano, Q

Exactive orbitrap, Thermo Scientific). The data analysis

was performed with Proteome Discoverer 1.4 by searching

against the sequences of cgAUS1 (A0A075DN54) and

cgAUS2 (A0A075BWS7). The search engine applied was

Sequest. The peptide mass tolerance was 5 ppm and the

fragment mass tolerance 0.5 Da. Carbamidomethylation of

cysteines was set as a static modification, whereas phos-

phorylation of serine, threonine and tyrosine as well as

sulfation of tyrosine were set as variable modifications. For

high confidence in the MS data, the false discovery rate

(FDR) of the peptide spectrum matches (PSM) was set to

\0.01 (Proteome Discoverer). The search results were

filtered with cross-correlation (XCorr) scores, set as 2.0,

2.5, 3.0 and 3.5 for peptides carrying 1, 2 and 3 or 4

charges, respectively.

Results

Protein extraction and removal of pigments

Petals from all developmental stages of flowers show AUS

activity (Kaintz et al. 2014). To obtain a high amount of

aurone synthase, 6 kg petal tissue (3 batches of 2 kg) of all

developmental stages was used as starting material. Pig-

ments were quantitatively removed from the crude extract

by a combination of repeated Triton X-114-induced ATPS,

ammonium sulfate precipitation and iterated PEG-4000-

induced ATPS. The addition of ammonium sulfate to the

crude extract containing Triton X-114 at the given pH

lowered the cloud point to temperatures (8–12 �C) that areless harsh for proteins in comparison to the temperature

induced ATPS at 37 �C (Sojo et al. 1998). The consecutive

addition of ammonium sulfate to 30 % saturation led to

further removal of pigments by precipitation. Pigments still

present were removed by PEG-4000-induced ATPS with

descending additions of PEG-4000 to avoid protein pre-

cipitation. A clear, beige to colorless enzyme solution was

finally obtained.

Measurement of enzyme activity during protein

purification

During the optimization of the purification procedure, the

chalcone butein (Fig. 1) and the flavonol fisetin (Jimenez

et al. 1998), both possessing two vicinal hydroxyl groups in

ring B, were used in spectrophotometrical assays of the

obtained fractions. No differences in substrate specificity

between different fractions were observed. However, some

fractions showed a sigmoid time course when fisetin was

used as a substrate indicating an allosteric activation

mechanism (Valero and Garcia-Carmona 1992). Fractions

showing this behavior were, therefore, considered to con-

tain portions of latent cgAUS. Modification of the enzy-

matic assays either by addition of 2.5 mM SDS or by

applying a pH of 3.5 lead to a steady-state reaction rate.

Protein purification by Fast Protein Liquid

Chromatography (FPLC)

The chromatogram obtained by cation exchange (CEX)

chromatography on SP-Sepharose FF showed broad ac-

tivity peaks (Fig. 2a). The occurrence of several active

forms made it necessary to perform 5 purification steps (6

for the latent form). Due to limited column capacities and

sample shelf life, several interlaced purification lines were

performed in parallel. Related forms of identical purifica-

tion stages were combined guided by activity and con-

ductivity of the eluted fractions. In total, seven different

samples containing significant amounts of cgAUS, in total

Planta (2015) 242:519–537 523

123

approximately 5.9 mg, (Table 1) were purified and char-

acterized. Samples 1–4 contained active cgAUS and sam-

ples 5–7 contained latent cgAUS. Latent (sample 5) and

cleaved latent forms (samples 6–7) were separated on the

stage of Mono Q from the active forms (samples 1–4)

(Fig. 2b). Example chromatograms of the purification on

SP-Sepharose FF, Mono Q and the polishing steps on

Mono S of the samples 1–5 are shown in Fig. 2.

Fig. 2 Typical chromatograms of the purification procedure of latent

and active cgAUS. Enzymatic assays for active cgAUS were

performed by monitoring the oxidation of 1 ml 50 lM fisetin in

125 mM sodium citrate pH 5.5 at 280 nm (Jimenez et al. 1998).

2.5 mM SDS were added to the reaction mixture for latent and

proteolytically cleaved latent cgAUS assays. One unit of enzyme was

defined as the amount that catalyzed the formation of 1 nmol of

oxidized fisetin per minute. a CEX chromatography using SP-

Sepharose FF, pH 5.0. b AEX chromatography using Mono Q, pH

8.5. Fractions containing latent, proteolytically cleaved latent and

active cgAUS are indicated below the chromatogram. c Polishing

CEX chromatography of active sample 1 using Mono S, pH 5.0.

d Polishing CEX chromatography of active samples 2 and 3 using

Mono S, pH 5.0. e Polishing CEX chromatography of active sample 4

using Mono S, pH 5.0. f Polishing CEX chromatography of latent

sample 5 using Mono S, pH 5.6

524 Planta (2015) 242:519–537

123

Gel electrophoresis and mass determination

by ESI–QTOF–MS

The gels showed two bands for active cgAUS samples after

reducing and three bands after non-reducing SDS-PAGE at

a molecular weight of approx. 37 kDa (Fig. 3a, b, lane-

numbers 1–4, Table 1). The occurrence of several bands is

caused by incomplete denaturation and incomplete reduc-

tion of the protein, evidenced by ESI–QTOF–MS mole-

cular mass determination (Fig. 4; Table 2, Online

Resources Fig. S2; Table S1). Incomplete denaturation is

also visible for the latent cgAUS samples under non-re-

ducing conditions. The latent cgAUS (sample 5) showed a

mass of 54 kDa and 59 kDa under non-reducing and re-

ducing conditions (Fig. 3a, b, lane-number 5). Interest-

ingly, the latent samples 6 and 7 showed masses of the

latent cgAUS under non-reducing conditions, but mole-

cular masses of the active cgAUS under reducing condi-

tions (Fig. 3a, b, lane-numbers 6 and 7, Table 1). An

additional band at about 19 kDa occurred under reducing

conditions, which indicates that the C-terminal domain is

bound to the main core by a disulfide bridge. This

hypothesis was supported by mass determination of active

cgAUS samples 1–4 (compare Table 1) by ESI–QTOF–MS

under reducing and non-reducing conditions. The obtained

mass spectra are shown in Fig. 4 and in Online Resource

Fig. S2. The mass spectrum of cgAUS sample 1 under non-

reducing conditions showed two major species exhibiting

masses of 41,559.0 and 41,639.0 Da, with an intensity ratio

of about 1.4:1 and a mass difference of 80.0 Da (Fig. 4a,

species A and B). Several peaks with significantly lower

intensities, each exhibiting mass differences of ap-

proximately 16 Da, were also present for the two main

peaks, indicating that the protein might contain several

oxidized residues. The mass difference of 80 Da may be

caused by either a phosphorylation or a sulfation. Under

reducing conditions species A was shifted by -1,523.4 Da

and species B by -1,523.3 Da (Fig. 4b, species C and D),

with a peptide of the corresponding mass (E1, E2) ap-

pearing. The monoisotopic mass matched the calculated

value of the sequence D438GVFTTPCDPEYAGG452 of

cgAUS1. This sequence is localized downstream of the

main core in the C-terminal region of the pro-enzyme

(Fig. 5a). The existence of a disulfide bridge connecting

Table 1 Yields, molecular masses (SDS-PAGE) and the kinetic classification of purified cgAUS samples. 6 kg frozen petal material was used as

starting material

Sample Amount (mg) M (kDa) (reducing SDS-PAGE) M (kDa) (non-reducing SDS-PAGE) Latent/active

1 2.45 37; 38 36; 37; 38 Active

2 0.56 37; 38 36; 37; 38 Active

3 0.37 37; 38 36; 37; 38 Active

4 1.25 37; 38 36; 37; 38 Active

5 0.87 59 65, 61, 54 Latent

6 0.25 38; 19 65, 54 Latent

7 0.12 37; 19 65, 54 Latent

Molecular masses were determined by reducing and non-reducing SDS-PAGE. Satellite bands exhibiting a shift to lower masses are found due to

incomplete denaturation and/or incomplete reduction of the applied samples

Fig. 3 SDS-PAGE analysis of purified cgAUS. About 4 lg total

protein was loaded on each lane. Lane-numbers in a and b correspond

to the described cgAUS samples within this article. Lane-number 1–4:

active cgAUS; lane-number 5: latent cgAUS; lane-numbers 6 and 7:

proteolytically cleaved latent cgAUS; M, molecular weight marker.

a SDS-PAGE (according to Laemmli 1970) of purified cgAUS.

b SDS-PAGE of purified cgAUS under semi-denaturating conditions:

Laemmli sample buffer without DTT, samples were not boiled

Planta (2015) 242:519–537 525

123

the main core of cgAUS1 with the shielding C-terminus is

thus supported by the ESI–QTOF–MS experiments.

Theoretical masses for the cgAUS1 sequence

(A0A075DN54) were calculated by taking into account 3

disulfide linkages (-6H), the thioether crosslink (-2H) as

well as the C-terminally hydrolyzed peptide backbone

(?H2O). Based on these calculations the primary structure

of the active cgAUS1 (sample 1) was deduced (Table 2;

Fig. 5a, the primary structure is highlighted by red

rectangles).

The mass spectra of active cgAUS samples 2–4 (Online

Resource Fig. S2) showed several peaks. The peaks that

differ in 80 Da from each other show intensity ratios

comparable to the observed ratios of sample 1. Calculations

Fig. 4 Positive mode ESI-

QTOF mass spectra of purified

active and latent cgAUS1

samples. Entire and magnified

mass spectra of acidified sample

1 containing active cgAUS1.

a Under non-reducing

conditions (untreated). b Under

reducing conditions

(preincubated with

50 mM DTT). c Mass spectra of

sample 5 containing acidified

latent cgAUS1 under non-

reducing conditions

526 Planta (2015) 242:519–537

123

of theoretical masses of cgAUS1 showed that the mass

differences between the different species are caused by a

combination of a phosphorylation/sulfation and unspecific

proteolytic cleavage occurring at the C-terminus of the

main core and at the remaining C-terminal peptide (Online

Recource Table S1). The terminal amino acids of these

species are indicated by a red color font in Fig. 5a.

The mass spectrum of latent cgAUS (sample 5) under

non-reducing conditions displayed two peaks with an in-

tensity ratio of 1:3.6 (Fig. 4c). Charge deconvolution

yielded masses of 58,928.2 Da, matching the mass of latent

cgAUS1, and 58,858.0 Da, respectively. The resulting

mass difference can be explained as a cleaved alanine,

most likely caused by proteolytic cleavage of A517

(Table 2; Fig. 5a).

Protein identification

Identification of the purified enzyme as cgAUS1

(A0A075DN54) was accomplished via HPLC–ESI–MS/

MS experiments on tryptic digests of latent cgAUS (sample

5, sequence coverage: 85 %, 441/517 amino acids) and

proteolytically activated cgAUS (sample 1, sequence cov-

erage: 77 %, 272/350 amino acids). The lists of the found

peptides are presented in Online Resource Table S2.

Only three unique peptides of the cgAUS2 sequence

(A0A075BWS7) were found in the tryptic digest of a so-

lution of sample 1. In the case of the latent sample 5 only

one unique peptide of cgAUS2 was found. Some peptides

of the C-terminal domain of cgAUS1 were found in the

active enzyme sample. This indicates that the active

cgAUS1 sample 1 contains trace amounts of latent enzyme.

cgAUS1 was found to be phosphorylated (Dmass:

-2.78 ppm, XCorr: 6.26, charge: 3) or sulfated (Dmass:

-3.56 ppm, XCorr: 6.45, charge: 3) at Tyr230. This

phosphorylation/sulfation was also found in the latent

sample 5. Whether this modification is really present in the

latent form or the observation is caused by trace amounts of

active cgAUS in the sample (compare Fig. 3a, b) remains

unclear.

Kinetic properties of active aurone synthase

Kinetic analysis of aurone synthase was performed on a

variety of substrates listed in Table 3 and included the two

60-deoxychalcones butein and marein, the 4-deoxyaurone

sulfuretin, the flavonol fisetin, and four common PPO

substrates. No differences in activity and substrate speci-

ficity were observed between the described active cgAUS

forms (samples 1–4, identified by HPLC–ESI–MS/MS and

ESI–QTOF–MS experiments as cgAUS1). The kinetic

properties of active cgAUS1 were determined using sample

1. To assess substrate specificity of cgAUS1 and a putative

specific involvement in aurone formation, purified vvCO

was used as a reference enzyme for a common catechol

oxidase.

Two subsequent reactions, catalyzed by cgAUS1, were

spectrophotometrically observed when butein was used as

a substrate (Fig. 6a, b). The first reaction (Fig. 6a) rep-

resents the oxidation of butein to sulfuretin and causes a

maximal increase in absorbance at 415 nm. This as-

sumption was verified by calculating difference spectra of

commercially available butein and sulfuretin

(Dkmax = 415 nm) as well as analysis of the reaction

products by means of RP-HPLC (Fig. 7). The following

mechanism for this reaction has been proposed for

AmAS1: butein is enzymatically oxidized to its chalcone

quinone, followed by an oxidative cyclization yielding

the aurone sulfuretin which is thought to take place non-

enzymatically (Nakayama et al. 2001).

Table 2 Masses determined from deconvoluted mass spectra of active and latent cgAUS1 under reducing and non-reducing conditions

Sample Species M (exp.) (Da) Deduced sequence M (calc.) [Da]

1 A 41,559.0 ± 0.3 API-IENSKE//DGVFTTPCDPEYAGG 41,559.35

B 41,639.0 ± 0.4 API-IENSKE ? X//DGVFTTPCDPEYAGG 41,639.35

1 reduced C 40,035.6 ± 0.3 API-IENSKE 40,036.79

D 40,115.7 ± 0.3 API-IENSKE ? X 40,116.79

E 764.8207 [M ? 2H]2? (mono)

1,528.6318 [M ? H]? (mono)

DGVFTTPCDPEYAGG 764.8167 [M ? 2H]2? (mono)

1,528.6261 [M ? H]? (mono)

5 F 58,928.2 ± 0.6 API-PIPKA 58,927.37

G 58,858.0 ± 1.7 API-PIPK 58,856.29

Theoretical masses were calculated by including the two disulfide bridges, one thioether-bridge [existence assumed by similarity to plant

catechol oxidases PPO_VITVI (P43311) and PPO1_IPOBA (Q9ZP19)], the previously unknown disulfide bond linking the C-terminus to the

main core as well as the C-terminally hydrolyzed peptide backbone

X is assumed to have a molecular weight of 80.0 Da

Planta (2015) 242:519–537 527

123

The second reaction (Fig. 6b) was observed only after

the majority of butein had been oxidized to sulfuretin. This

was visible by a decrease in absorbance at 415 nm attended

by a shift to higher wavelengths and a newly occurring

broad absorption band from 480–600 nm. Sulfuretin is an

o-diphenol and therefore a potential substrate for PPOs.

The time course of the oxidation of sulfuretin

(Dkmax = 475 nm) showed a nonlinear behavior with a

slope decreasing over time even reaching an idle state,

although only a minority of substrate has been oxidized

(Fig. 6c). The final quantity of product formation strongly

depended on the amount of enzyme used for the assay

(Fig. 6c). The observed decrease in absorbance is most

likely a consequence of polymerization of the oxidation

products indicated by the broad absorption band around

520 nm and also indicated by HPLC analysis from assays

528 Planta (2015) 242:519–537

123

with high amounts of protein, in which neither products nor

substrates could be observed.

In contrast to AmAS1, no hydroxylation reaction was

detected when isoliquiritigenin was used as a substrate,

even when hydrogen peroxide was added to the reaction

medium to omit the lag phase by transferring the copper

center of the protein to the reactive oxy-form (reviewed by

Ramsden and Riley 2014). In addition, no introduction of a

third hydroxyl group in the B-ring of the substrates butein

and marein could be observed by means of HPLC analysis.

The pH optimum of cgAUS1 is between pH 5.4–6.5 for

butein (Fig. 6d). Due to the higher stability of o-diphenolic

substrates (as well as the resulting o-quinones) at acidic pH

values, kinetic parameters for cgAUS1 as well as for vvCO

were determined at pH 5.5. Marein and fisetin showed a

low affinity to cgAUS1 and the expected Km values were

significant higher than the solubility of these substrates (up

to 75 lM). This prevented an accurate estimation of Km by

nonlinear regression. Instead kcat/Km values were deter-

mined by linear fitting to pseudo first-order kinetics

([S] � Km). This was also the case for butein, marein and

fisetin with vvCO enzyme. cgAUS1 displayed a Km value

of 52 ± 3 lM and also the highest catalytic efficiency

towards butein (Table 3). The kcat/Km value of marein is

more than five times lower than for butein. vvCO instead

showed the highest catalytic efficiency towards fisetin and

a more than six times lower efficiency towards butein than

cgAUS1. In general, cgAUS1 exhibits lower Km values and

possesses higher kcat/Km values. 4-Methylcatchol is the

only exception showing very similar catalytic efficiencies

for both enzymes.

Kinetic properties of latent aurone synthase

The recombinantly expressed cgAUS1 (Kaintz et al. 2014)

exhibited identical properties to the latent cgAUS1 purified

from the natural source. However, the enzyme purified

from Coreopsis petals contained low amounts of active

cgAUS1. Recombinantly expressed cgAUS1 was therefore

used to describe the characteristics of the latent enzyme.

The pH optimum of latent cgAUS1 reflects the acidic

activation of the pro-enzyme in a very narrow range around

pH 4.0 (Fig. 6d). An allosteric activation was observed

when fisetin was used at pH 5.5 as a substrate similar to

that described for latent vvCO (Valero and Garcia-Car-

mona 1992). The lag period is dependent on the enzyme

concentration but the steady-state rate is reached at a dis-

tinct product concentration (Fig. 6e). An in situ product

concentration of approximately 23 lM is determined to be

the critical concentration in the case of fisetin. At this

concentration of oxidized fisetin cgAUS1 is fully activated.

The experiments also show that a potential influence of the

incubation time of the enzyme with oxidized fisetin is

negligible under the chosen conditions. Allosteric activa-

tion was observed for all substrates tested, except for

chalcones where no activiation occurred. Instead, cgAUS1

showed a steady-state reaction rate, but only a few percent

of its maximum activity at pH 4.0 during the complete

reaction time (comp. Figure 6d).

Discussion

Purification and identification of aurone synthase

Polyphenols and their corresponding quinones are highly

reactive compounds that form polymerized products. These

are responsible for protein cross-linking and cause protein

precipitation (Mcmanus et al. 1981; Ito et al. 1984;

McDowell et al. 1999; Kumar et al. 2000). It is absolutely

essential therefore, to remove these compounds from crude

extracts which serve as a source for a pure protein. ATPS is

a commonly used first step in protein isolation/purification

(reviewed by Aguilar and Rito-Palomares 2010 and Hong

Yang 2013). In this study ATPS systems based on Triton

X-114 and PEG (Sojo et al. 1998) were combined to

bFig. 5 Sequence analysis and phylogenetic tree of cgAUS1 in

comparison with other PPOs. a Primary structure of cgAUS1

excluding the transit peptide. Red rectangles are used in the alignment

to highlight the deduced primary structure of the sample 1 whereas

terminal amino acids of species of minor abundance (compare ESI-Q-

TOF measurements, Fig. S2; Table S1) are written in a red color font.

Copper-binding histidines are highlighted blue, copper-binding sites

are underlined with blue dashed lines. Proteolytic cleavage sites are

marked by violet lines. Disulfide linkages and the thioether-bridge are

marked by yellow connectors. The phosphorylated/sulfated tyrosine

residue (identified by HPLC–ESI–MS/MS experiments) is highlighted

in orange. The insertion in the loop region near the active site of

catechol oxidases (PPO_VITVI pdb-entry: 2P3X; PPO1_IPOBA pdb-

entry: 1BT3) is colored in green and the highly conserved KFDV-

motif is marked by a green rectangle. b Sequence alignments of CuB

binding sites of several plant PPOs (monophenolase/diphenolase

activity: (a) Klabunde et al. 1998, (b) Virador et al. 2010, (c) Escobar

et al. 2008; Zekiri et al. 2014, (d) Cho et al. 2003, (e) Nakayama et al.

2000, (f) Dirks-Hofmeister et al. 2012, (g) this work). The copper-

binding histidines are highlighted blue, the conserved phenylalanine

above the active site is highlighted in pale blue, the insertion of group

2 PPOs in the loop region near the active site is highlighted in green.

For clarity the sequences of dandelion PPO-9 and PPO-11 were

omitted, because the insertion is even larger. The positions HB1?1

and HB2?1 are marked by red rectangles. c Phylogenetic tree of

several plant PPOs. The corresponding alignment of these PPOs is

presented in Online Resource Fig. S3. d Sequence alignments near the

C-terminal residual peptide found to be connected by a disulfide

linkage to the main core of mature cgAUS1. The KFDV-motif is

highlighted with green rectangles and the cysteines are shaded in

yellow. The primary structure of cgAUS1 and sequence alignments

were prepared using ALINE (Bond and Schuttelkopf 2009). The

phylogenetic tree was prepared using TreeGraph 2 (Stover and Muller

2010)

Planta (2015) 242:519–537 529

123

remove pigments and polyphenols which accumulate in the

more hydrophobic detergent-rich phases. The successful

combination of Triton X-114 ATPS, ammonium sulfate

precipitation and PEG-4000 ATPS is a decisive improve-

ment in the protein purification procedure from polyphenol

rich natural sources.

The chromatograms, especially at a small scale, dis-

played good resolutions of differing protein peaks. AUS

activity, however, could be determined over a wide range

of the gradient. This indicates that cgAUS might contain

diverse modifications, resulting in overlapping peaks in the

chromatograms. Cation and anion exchange chromatogra-

phy were therefore combined to benefit from different ti-

tration curves of the different forms. From the large

number of AUS peaks in the obtained chromatograms,

those containing high activity and high amounts of cgAUS

were chosen for further purification and investigation.

Despite the diversity and high number of active cgAUS

forms observed during purification, only one latent and two

partially proteolytically cleaved forms of lower quantity

were observed during purification. Mass spectrometric

analyses revealed that all the active cgAUS forms were

caused by unspecific proteolytic cleavage of cgAUS1 in

combination with a phosphorylation or sulfation. Recent

works also reported an unspecific C-terminal proteolytic

cleavage of PPOs resulting in several heterogeneous forms

(Mauracher et al. 2014; Zekiri et al. 2014). Only a small

number of unique peptides from cgAUS2 were found. The

abundance and prevalence of cgAUS1 in petals of C.

grandiflora is in accordance with reported gene expression

studies (Kaintz et al. 2014) which also showed a correlation

of cgAUS1 expression with aurone accumulation in petals

of C. grandiflora.

Unique structural features of cgAUS1, a member

of the novel group 2 PPOs

cgAUS1 shows the highest sequence identity for the cat-

alytically active domains (53–65 %, excluding the transit

peptide and the C-terminal domain) with PPOs from dan-

delion (Taraxacum officinale) and a considerably lower

sequence identity to other PPOs (40–50 %). It was sug-

gested recently, that dandelion PPOs are clustered into two

distinct groups (Dirks-Hofmeister et al. 2014). The

grouping of these PPOs correlates with differences at the

CuB binding site, namely at the amino acid directly

Table 3 Kinetic parameters of cgAUS1 and vvCO

Substrate k (nm) De (l mol-1 cm-1) Km (mM) Vmax (lmol min-1 mg-1) kcat (s-1) kcat/Km (mM s-1)

Aurone synthase (Coreopsis grandiflora)

Butein 415 9,320a 0.052 ± 0.003 489 ± 17 464 ± 16 8,927 ± 649

Marein 425 13,559b –c –c –c 1,647 ± 62c

Sulfuretin 475 –d –d –d –d –d

Fisetin 280 5,345e –c –c –c 2,130 ± 39c

4-tert-Butylcatechol 400 1,150f 0.504 ± 0.020 2,256 ± 23 1,563 ± 16 3,101 ± 128

Chlorogenic acid 400 2,566g 0.984 ± 0.080 2,549 ± 85 1,782 ± 63 1,811 ± 160

4-Methylcatechol 400 1,350f 3.11 ± 0.12 2,299 ± 39 1,593 ± 27 512 ± 22

Catechol 390 1,450f 4.04 ± 0.69 1,255 ± 59 869 ± 41 215 ± 38

Catechol oxidase (Vitis vinifera)

Butein 415 9,320a –c –c –c 1,300 ± 32c

Marein 425 13,559b –c –c –c 472 ± 8c

Sulfuretin 475 –d –d –d –d –d

Fisetin 280 5,345e –c –c –c 1,902 ± 35c

4-tert-Butylcatechol 400 1,150f 2.43 ± 0.19 4,110 ± 110 2,847 ± 76 1,170 ± 98

Chlorogenic acid 400 2,566g 2.00 ± 0.18 2,274 ± 86 1,575 ± 59 786 ± 76

4-Methylcatechol 400 1,350f 7.10 ± 0.12 5,320 ± 33 3,685 ± 22 519 ± 9

Catechol 390 1,450f 45.88 ± 1.67 2,857 ± 41 1,979 ± 28 43.1 ± 1.7

a Extinction coefficient was determined by calculating difference spectra of sulfuretin and buteinb Extinction coefficient was determined at the maximum increase of absorbance by assuming 100 % product formationc A pseudo first-order kinetic model ([S] � Km) was used to calculate the ratio kcat/Km due to low solubility of the substratesd Extinction coefficient could not be determined due to complex and unspecific product formatione (Jimenez et al. 1998)f (Waite 1976)g (Munoz et al. 2007)

530 Planta (2015) 242:519–537

123

following the second copper-binding histidine (HB2 ? 1)

(Dirks-Hofmeister et al. 2014). This residue is a bulky,

charged arginine in group 2 PPOs and a small uncharged

residue in group 1 PPOs. Notably, the selected PPOs ex-

hibiting monophenolase activity share small and hy-

drophobic residues in this position (Fig. 5b). Crystal

structures of tyrosinase from Bacillus megaterium

demonstrated the importance of residue HB2 ? 1 for

docking of substrates to the active site (Goldfeder et al.

2014) recently. Due to the absence of a charged residue in

this position an alternative substrate docking for plant ty-

rosinases seems to be very likely.

Phylogenetic analysis shows that cgAUS1 clusters in the

group 2 PPOs from T. officinale (Fig. 5c, the corresponding

Fig. 6 Enzymatic properties of latent and active cgAUS1. a,b Difference spectra of the enzymatic oxidation of butein by

cgAUS1. The reaction medium contained 25 lM butein in

125 mM sodium citrate buffer, pH 5.5. The final concentration of

active cgAUS1 was 0.9 nM. Spectra were recorded every 2 min.

a Oxidative conversion of butein to sulfuretin (0–12 min). b Subse-

quent oxidation of sulfuretin (compare Fig. 8). c Time course of the

oxidation of sulfuretin by cgAUS1 monitored at 475 nm indicating

suicide inactivation. The reaction medium contained 50 lM sulfuretin

in 125 mM sodium citrate buffer, pH 5.5. The final concentrations of

active cgAUS1 were 1.2 nM (1), 0.6 nM (2), 0.3 nM (3) and

0.15 nM (4). d pH optimum of recombinant pro-cgAUS1 (filled

triangle, dashed lines) and active cgAUS1 (filled circle, solid lines).

25 lM Butein in 125 mM sodium citrate buffers was used over a range

from pH 3.5 to 7.4. e Allosteric activation of latent recombinant

cgAUS1 monitored at 282 nm. The reaction medium contained

50 lM fisetin in 125 mM sodium citrate buffer, pH 5.5. The final

concentrations of latent recombinant cgAUS1 were 7.2 nM (1),

3.6 nM (2), 1.8 nM (3) and 0.9 nM (4). The dashed line depicts the

beginning of the steady-state region. The absorbance of 120 mAU

corresponds to a product concentration of 23 lM

Planta (2015) 242:519–537 531

123

alignment is presented in Online Resource Fig. S3).

Therefore we propose that these PPOs form a novel and,

with exception of recombinantly expressed pro-PPOs from

T. officinale, previously uncharacterized subclass of plant

PPOs. The sequence alignment of group 1 and group 2

PPOs expose a novel characteristic feature of the group 2

PPOs (Fig. 5b). There exists an insertion just before the

CuB binding site (Fig. 5b, e.g. V237ANG240 in the cgAUS1

sequence). Due to its location in a loop region on the

surface and near to the active site of catechol oxidases

(pdb-entries: 1bt3, 2p3x) it might influence substrate

docking to the active site. This might also explain the more

specific substrate binding of cgAUS1 as cgAUS1 displays

generally lower Km values and, with exception of fisetin, a

higher catalytic efficiency than vvCO (Table 3).

Indications of a putative phosphorylated or sulfated

residue of active cgAUS1 forms were obtained by ESI–

QTOF–MS experiments. The residue Tyr230 was found to

be phosphorylated or sulfated by means of HPLC–ESI–

MS/MS experiments of tryptic digested active cgAUS1

with a high degree of confidence (Online Resource Table

S2). To date, only one PPO from dormant terminal buds in

poplar (Populus simonii 9 P. nigra) was found to be

phosphorylated (Liu et al. 2011). The phosphorylation in

poplar PPO, however, was found in the shielding C-ter-

minal domain and not in the main core. Notably, the

Fig. 7 HPLC chromatogramms of the chalcone butein, the aurone sulfuretin and after incubation with active cgAUS1. a Butein without

incubation treatment. b Sulfuretin without incubation treatment. c Butein incubated with cgAUS1

532 Planta (2015) 242:519–537

123

modification in cgAUS1 was found in the immediate

vicinity of the loop extension (Fig. 5a). Its role remains

unclear at this stage of the work.

The results of analytical SDS-PAGE for the purified

latent cgAUS forms differ under reducing and non-reduc-

ing conditions. Consequently, we concluded that the

C-terminal domain, shielding the active site of the protein,

is connected to the main core of the enzyme by a disulfide

bond. This hypothesis was verified by ESI–QTOF–MS

measurements of the active cgAUS forms (sample 1 to

sample 4) under reducing and non-reducing conditions.

The residual C-terminal peptide of active cgAUS starts a

few amino acids behind the KFDV-motif, which is highly

conserved in plant PPOs (Fig. 5d). We assume, that the

main core cysteine Cys206 is involved in disulfide linkage,

as the highly conserved Cys334 is not accessible (comp.

pdb-entry 2p3x). An intermolecular disulfide linkage has

been reported to be responsible for the tetramerization of

recombinant expressed PPO-6 and PPO-7 from T. officinale

(Dirks-Hofmeister et al. 2012, 2014) and Cys197 was

identified to be involved in the tetramerization. An in-

tramolecular crosslink between the catalytically active

main core and the shielding C-terminal domain has not

been reported so far and one function might be that it

stabilizes the interface between the two domains. The pro-

enzyme has to be cleaved at three different positions to

result in the active form (Fig. 5a, cleavage sites are indi-

cated by violet lines). An involvement of the remaining

C-terminal peptide (D438GVFTTPCDPEYAGG452) in the

catalytic activity of the enzyme is very unlikely, because

the peptide is short (7 amino acids upstream and down-

stream of Cys445, respectively), and the Cys206 is located in

a helix further away of the active site.

Kinetic characterization suggests alternative

4-deoxyaurone formation in Asteraceae species

In the assays with purified cgAUS1, the oxidation of

butein to sulfuretin followed the route described for

AmAS1 (Fig. 6a). Additionally, the oxidation of sul-

furetin to the corresponding o-quinone could be observed

as well (Fig. 6b). However, this was strongly dependent

on the presence of large enzyme amounts in the assay

(Fig. 6c) and can be explained by the suicide inactivation

of PPOs (Escribano et al. 1989; Chazarra et al. 1997;

Garcia-Molina et al. 2005; Land et al. 2008; Munoz-

Munoz et al. 2008, 2011, 2012a, b; Ramsden et al. 2009;

Ramsden and Riley 2010a). It has been reported that

AmAS1 shows virtually no oxidation of aurones

(Nakayama et al. 2000) and comparison with our results

suggests that AmAS1 might exhibit suicide inactivation

towards aurones. In accordance to literature (Chazarra

et al. 1997; Ramsden and Riley 2010b), the suicide

substrates 4-tert-butylcatechol and 4-methylcatechol were

identified to inactivate cgAUS1. As expected, vvCO

demonstrated irreversible inactivation towards p-substi-

tuted substrates as well.

Kinetic parameters demonstrate a high specificity and

efficiency of cgAUS1 for butein in comparison to vvCO.

As butein is a naturally occurring substrate in Coreopsis

species (Crawford and Smith 1983), this supports the as-

sumption that cgAUS1 is specifically involved in the au-

rone biosynthesis. The significant higher affinity of butein

to cgAUS1 than to vvCO indicates that specific residues

of cgAUS1 are involved in the coordination of butein to

its active site. Confirmation and identification of potential

residues are, however, subject of analysis of crystal

structures of active cgAUS1. In contrast to the determi-

nation of kinetic parameters of cgAUS1 towards butein

(nonlinear regression), the kcat/Km value of cgAUS1 to-

wards marein had to be determined by linear fitting to

pseudo first-order kinetics ([S] � Km), although both

substrates (butein and marein) were applied at the same

concentration ranges (10–75 lM). This indicates a sig-

nificantly lower affinity of cgAUS1 to marein. Unfortu-

nately, to date, no chalcone glycoside/chalcone aglycone

pair is commercial available, therefore it remains unclear

whether this difference in affinity is caused by the gly-

cosyl group of marein or by the hydroxyl group at posi-

tion 30. However, the occurence of chalcone aglycones

(butein, okanin, lanceoletin) and the corresponding aurone

aglycones (sulfuretin, maritimetin, leptosidin) in Coreop-

sis (comp. Fig. 1) combined with the observed lower

affinity of marein strongly support that 4-deoxyaurone

formation in C. grandiflora occurs at the level of chalcone

aglycones possessing two vicinal hydroxyl groups in the

B-ring. This hypothesis is furthermore supported by the

reported significantly higher affinity of a glycosyltrans-

ferase from C. grandiflora for sulfuretin than for butein

(Halbwirth et al. 1997), indicating that glycosylation oc-

curs on the level of aurones. Combination with previous

studies on 4-deoxyaurone formation in Bidens ferulifolia

(Miosic et al. 2013) strongly suggests that an alternative

pathway for aurone biosynthesis exists in Asteraceae

species, in addition to that proposed for A. majus (Ono

et al. 2006a) (Fig. 8). The predicted localization in

chloroplasts (or chromoplasts) of cgAUS1 (Kaintz et al.

2014) does not contradict its involvement in aurone

biosynthesis. Recently, Araji et al. (2014) proposed a

novel involvement of PPOs, most likely localized in

chloroplasts, in the secondary metabolism of walnut

(Juglans regia) leaves. The authors suggested that the

substrates, biosynthesized in the cytoplasm, would pre-

sumably need to be transported back to the chloroplast for

catalysis by PPO. However, to date no information of

prerequisite transport mechanisms are available.

Planta (2015) 242:519–537 533

123

Allosteric activation: latent cgAUS1 is fully activated

at a distinct o-quinone concentration

Although allosteric activation of latent PPOs has been re-

ported some time ago and several latent PPOs are recom-

binantly available, the phenomenon is rarely described and

still poorly understood. Allosteric activation of cgAUS1

occurs at neutral or slightly acidic pH ranges. This be-

havior has also been reported for other latent PPOs (Valero

and Garcia-Carmona 1992, 1998; Nillius et al. 2008; Dirks-

Hofmeister et al. 2012, 2014). Previous studies reported

that the lag period remained constant with varying the

enzyme concentration (Valero and Garcia-Carmona 1992).

In the case of latent cgAUS1, however, the lag period

strongly depended on enzyme concentration and the steady

state was reached at a defined product concentration

(Fig. 6e). The absence of allosteric activation during the

oxidation of chalcones suggests that the activation of latent

cgAUS1 is caused by o-quinones, as aurones themselves do

not possess a quinoide structure. The quinone-binding site

might be within the interface of the shielding C-terminal

domain and the main core, which might result in a similar

structural rearrangement as proposed for an acidic activa-

tion or activation with SDS. The interaction is most likely

irreversible and covalent, because the o-quinones are

known to be highly reactive and likely to undergo rapid

polymerization reactions and are also responsible for pro-

tein cross-linking (Ito et al. 1984; Burzio and Waite 2000;

Rollett et al. 2013). The existence of a concentration

threshold of o-quinones where the enzyme is fully acti-

vated is a novel finding. However, the principle behind this

behavior remains unclear and may vary, depending on the

reactivity of the o-quinone. We propose that allosteric ac-

tivation can be a general feature of pro-PPOs and may

trigger important physiological defense mechanisms, for

example, those induced by tissue damage or during ox-

idative stress in plants and also in fungi.

Conclusion

The purified cgAUS was identified as cgAUS1 and the

corresponding cDNA clone was reported to be particularly

expressed in petals recently. Our results reveal that aurone

Fig. 8 4-Hydroxyaurone and proposed 4-deoxyaurone biosynthetic

pathway. The upper part shows the aurone biosynthetic pathway in

Antirrhinum majus (4-hydroxyaurone formation). Chalcones are

glycosylated by a chalcone 40-O-glucosyltransferase (40CGT) and

transported to the vacuole. Aureusidin synthase (AmAS1) possesses

monophenolase and diphenolase activity. The lower figure shows the

proposed biosynthetic pathway in C. grandiflora (4-deoxyaurone

formation). Chalcones are hydroxylated by a chalcone 3-hydroxylase

(CH3H); aurone synthase (cgAUS1) possesses only diphenolase

activity; aurones are glycosylated by a glycosyltransferase. Oxidation

of aurones resulted in suicide inactivated enzyme in vitro

534 Planta (2015) 242:519–537

123

synthase is a member of the novel group 2 PPOs and that

an insertion in a loop region near to the active site, which

might be involved in substrate docking, is characteristic for

this group. A phosphorylation/sulfation of unknown func-

tion was found in the immediate vicinity of the loop ex-

tension. The disulfide crosslink of the C-terminal domain to

the main core is a novel structural feature of plant PPOs.

Kinetic characterization of cgAUS1 suggests that aurone

formation might occur at the stage of chalcone agylcones

which would constitute an alternative aurone biosynthetic

pathway in Asteraceae species in comparison to that de-

scribed for A. majus (Plantaginaceae). The availability of

large amounts of highly purified and characterized aurone

synthase enables crystallization experiments. Structural

analysis of latent and active cgAUS1 will provide further

insights to complement the presented novel findings.

Author contribution CM.: Planned experiments; Per-

formed experiments; Analyzed data; Wrote the paper;

S.G.M.: Planned experiments; Analyzed data; S.P.: Per-

formed experiments; R.L.M.: Performed experiments;

Analyzed data; H.H.: Contributed reagents or other

essential material; Performed experiments; Analyzed data;

other; A.R.: Planned experiments; Contributed reagents or

other essential material; other.

Acknowledgments The research was funded by the Austrian Sci-

ence Fund (FWF): P25217-N28. Annette Rompel is grateful for fi-

nancial support by the Deutsche Forschungsgemeinschaft (DFG) Ro

1084/8-1. Heidi Halbwirth acknowledges funding by FWF (P24331-

B16). We thank Florime Zekiri for practical help during the protein

isolation procedure. We acknowledge Prof. Andreas Rizzi and

Claudia Michael for support during ESI–QTOF–MS experiments. We

thank Silvija Miosic for support in the performance of the HPLC

enzyme assays. Thanks for cultivating C. grandiflora and for taking

care of the plant fields go to the horticultural department of Molecular

Systems Biology, UZA1-Glashaus1, University of Vienna, 1090 Vi-

enna, Austria especially to Thomas Joch and Andreas Schrofl and the

gardeners of experimental garden Augarten, 1020 Vienna, Austria

especially to Miroslav Crep and Erich Wagner.

Open Access This article is distributed under the terms of the

Creative Commons Attribution License which permits any use, dis-

tribution, and reproduction in any medium, provided the original

author(s) and the source are credited.

References

Aguilar O, Rito-Palomares M (2010) Aqueous two-phase systems

strategies for the recovery and characterization of biological

products from plants. J Sci Food Agric 90:1385–1392. doi:10.

1002/jsfa.3956

Araji S, Grammer TA, Gertzen R, Anderson SD, Mikulic-Petkovsek

M, Veberic R, Phu ML, Solar A, Leslie CA, Dandekar AM,

Escobar MA (2014) Novel roles for the polyphenol oxidase

enzyme in secondary metabolism and the regulation of cell death

in walnut. Plant Physiol 164:1191–1203. doi:10.1104/pp.113.

228593

Bond CS, Schuttelkopf AW (2009) ALINE: a WYSIWYG protein-

sequence alignment editor for publication-quality alignments.

Acta Crystallogr D Biol Crystallogr 65:510–512. doi:10.1107/

S0907444909007835

Burzio LA, Waite JH (2000) Cross-linking in adhesive quinoproteins:

studies with model decapeptides. Biochemistry

39:11147–11153. doi:10.1021/bi0002434

Chandra A, Rana J, Li Y (2001) Separation, identification, quantifi-

cation, and method validation of anthocyanins in botanical

supplement raw materials by HPLC and HPLC-MS. J Agric

Food Chem 49:3515–3521. doi:10.1021/jf010389p

Chazarra S, Cabanes J, Escribano J, Garcia-Carmona F (1997) Kinetic

study of the suicide inactivation of latent polyphenoloxidase

from iceberg lettuce (Lactuca sativa) induced by 4-tert-butyl-

catechol in the presence of SDS. Biochim Biophys Acta

1339:297–303. doi:10.1016/S0167-4838(97)00012-5

Cho M-H, Moinuddin SGA, Helms GL, Hishiyama S, Eichinger D,

Davin LB, Lewis NG (2003) (?)-Larreatricin hydroxylase, an

enantio-specific polyphenol oxidase from the creosote bush

(Larrea tridentata). Proc Natl Acad Sci USA 100:10641–10646.

doi:10.1073/pnas.1934562100

Crawford DJ (1970) Systematic studies on mexican Coreopsis

(Compositae). Coreopsis mutica: flavonoid chemistry, chromo-

some numbers, morphology, and hybridization. Brittonia

22:93–111. doi:10.2307/2805804

Crawford DJ, Smith EB (1980) Flavonoid chemistry of C. grandiflora

(Compositae). Brittonia 32:154–159. doi:10.2307/2806783

Crawford DJ, Smith EB (1983) The distribution of anthochlor floral

pigments in north-american Coreopsis (Compositae): taxonomic

and phyletic interpretations. Am J Bot 70:355–362. doi:10.2307/

2443243

Davies KM, Marshall GB, Bradley JM, Schwinn KE, Bloor SJ,

Winefield CS, Martin CR (2006) Characterisation of aurone

biosynthesis in Antirrhinum majus. Physiol Plant 128:593–603.

doi:10.1111/j.1399-3054.2006.00774.x

Dirks-Hofmeister ME, Inlow JK, Moerschbacher BM (2012) Site-

directed mutagenesis of a tetrameric dandelion polyphenol

oxidase (PPO-6) reveals the site of subunit interaction. Plant

Mol Biol 80:203–217. doi:10.1007/s11103-012-9943-9

Dirks-Hofmeister ME, Singh R, Leufken CM, Inlow JK, Moer-

schbacher BM (2014) Structural diversity in the dandelion

(Taraxacum officinale) polyphenol oxidase family results in

different responses to model substrates. PLoS One 9:e99759.

doi:10.1371/journal.pone.0099759

Escobar MA, Shilling A, Higgins P, Uratsu SL, Dandekar AM (2008)

Characterization of polyphenol oxidase from walnut. J Am Soc

Hort Sci 133:852–858

Escribano J, Tudela J, Garciacarmona F, Garciacanovas F (1989) A

kinetic-study of the suicide inactivation of an enzyme measured

through coupling reactions. Application to the suicide inactiva-

tion of tyrosinase. Biochem J 262:597–603

Espin JC, van Leeuwen J, Wichers HJ (1999) Kinetic study of the

activation process of a latent mushroom (Agaricus bisporus)

tyrosinase by serine proteases. J Agric Food Chem

47:3509–3517. doi:10.1021/jf9813539

Flurkey WH, Inlow JK (2008) Proteolytic processing of polyphenol

oxidase from plants and fungi. J Inorg Biochem 102:2160–2170.

doi:10.1016/j.jinorgbio.2008.08.007

Gandia-Herrero F, Jimenez-Atienzar M, Cabanes J, Garcia-Carmona

F, Escribano J (2005) Evidence for a common regulation in the

activation of a polyphenol oxidase by trypsin and sodium

dodecyl sulfate. Biol Chem 386:601–607. doi:10.1515/BC.2005.

070

Planta (2015) 242:519–537 535

123

Garcia-Molina F, Hiner AN, Fenoll LG, Rodriguez-Lopez JN, Garcia-

Ruiz PA, Garcia-Canovas F, Tudela J (2005) Mushroom

tyrosinase: catalase activity, inhibition, and suicide inactivation.

J Agric Food Chem 53:3702–3709. doi:10.1021/jf048340h

Geissman TA, Heaton CD (1943) Anthochlor pigments. IV. The

pigments of C. grandiflora Nutt. I. J Am Chem Soc 65:677–683.

doi:10.1021/Ja01244a050

Geissman TA, Heaton CD (1944) Anthochlor pigments. V. The

pigments of C. grandiflora Nutt. II. J Am Chem Soc 66:486–487.

doi:10.1021/Ja01231a056

Geissman TA, Moje W (1951) Anthochlor pigments. VIII. The

pigments of C. grandiflora Nutt. III. J Am Chem Soc

73:5765–5768. doi:10.1021/ja01156a080

Geissman TA, Harborne JB, Seikel MK (1956) Anthochlor pigments. XI.

The constituents of Coreopsis maritima. Reinvestigation of Coreop-

sis gigantea. J AmChemSoc 78:825–829. doi:10.1021/ja01585a032

Goldfeder M, Kanteev M, Isaschar-Ovdat S, Adir N, Fishman A

(2014) Determination of tyrosinase substrate-binding modes

reveals mechanistic differences between type-3 copper proteins.

Nat Commun 5:4505. doi:10.1038/ncomms5505

Halbwirth H, Wimmer G, Wurst F, Forkmann G, Stich K (1997)

Enzymatic glucosylation of 4-deoxyaurones and 60-deoxychal-cones with enzyme extracts of C. grandiflora Nutt. I. Plant Sci

122:125–131. doi:10.1016/s0168-9452(96)04553-0

Harborne JB (1967) Comparative biochemistry of the flavonoids.

Academic Press, New York

Harborne JB, Geissman TA (1956) Anthochlor pigments. XII.

Maritimein and marein. J Am Chem Soc 78:829–832. doi:10.

1021/ja01585a033

Hong Yang AMG (2013) Aqueous two-phase extraction advances for

bioseparation. J Bioprocessing & Biotechniques. doi:10.4172/

2155-9821.1000140

Ito S, Kato T, Shinpo K, Fujita K (1984) Oxidation of tyrosine

residues in proteins by tyrosinase. Formation of protein-bonded

3,4-dihydroxyphenylalanine and 5-S-cysteinyl-3,4-dihydrox-

yphenylalanine. Biochem J 222:407–411

Jimenez M, Escribano-Cebrian J, Garcia-Carmona F (1998) Oxida-

tion of the flavonol fisetin by polyphenol oxidase. Biochim

Biophys Acta 1425:534–542

Kaintz C, Molitor C, Thill J, Kampatsikas I, Michael C, Halbwirth H,

Rompel A (2014) Cloning and functional expression in E. coli of

a polyphenol oxidase transcript from C. grandiflora involved in

aurone formation. FEBS Lett 588:3417–3426. doi:10.1016/j.

febslet.2014.07.034

King RS, Flurkey WH (1987) Effects of limited proteolysis on broad

bean polyphenoloxidase. J Sci Food Agric 41:231–240. doi:10.

1002/jsfa.2740410305

Klabunde T, Eicken C, Sacchettini JC, Krebs B (1998) Crystal

structure of a plant catechol oxidase containing a dicopper

center. Nat Struct Biol 5:1084–1090. doi:10.1038/4193

Kumar G, Bristow JF, Smith PJ, Payne GF (2000) Enzymatic gelation

of the natural polymer chitosan. Polymer 41:2157–2168. doi:10.

1016/S0032-3861(99)00360-2

Laemmli UK (1970) Cleavage of structural proteins during the

assembly of the head of bacteriophage T4. Nature 227:680–685.

doi:10.1038/227680a0

Land EJ, Ramsden CA, Riley PA, Stratford MRL (2008) Studies of

para-quinomethane formation during the tyrosinase-catalyzed

oxidation of 4-alkylcatechols. Arkivoc 2:258–267

Liu CC, Liu CF, Wang HX, Shen ZY, Yang CP, Wei ZG (2011)

Identification and analysis of phosphorylation status of proteins

in dormant terminal buds of poplar. BMC Plant Biol 11:158.

doi:10.1186/1471-2229-11-158

Marquardt DW (1963) An algorithm for least-squares estimation of

nonlinear parameters. J Soc Ind Appl Math 11:431–441. doi:10.

2307/2098941

Marusek CM, Trobaugh NM, Flurkey WH, Inlow JK (2006)

Comparative analysis of polyphenol oxidase from plant and

fungal species. J Inorg Biochem 100:108–123. doi:10.1016/j.

jinorgbio.2005.10.008

Matern U, Reichenbach C, Heller W (1986) Efficient uptake of

flavonoids into parsley (Petroselinum hortense) vacuoles re-

quires acylated glycosides. Planta 167:183–189. doi:10.1007/

BF00391413

Mauracher SG, Molitor C, Michael C, Kragl M, Rizzi A, Rompel A

(2014) High level protein-purification allows the unambiguous

polypeptide determination of latent isoform PPO4 of mushroom

tyrosinase. Phytochemistry 99:14–25. doi:10.1016/j.phytochem.

2013.12.016

Mayer AM (2006) Polyphenol oxidases in plants and fungi: going

places? A review. Phytochemistry 67:2318–2331. doi:10.1016/j.

phytochem.2006.08.006

McDowell LM, Burzio LA, Waite JH, Schaefer J (1999) Rotational

echo double resonance detection of cross-links formed in mussel

byssus under high-flow stress. J Biol Chem 274:20293–20295.

doi:10.1074/jbc.274.29.20293

Mcmanus JP, Davis KG, Lilley TH, Haslam E (1981) The association of

proteins with polyphenols. J Chem Soc Chem Comm 7:309–311

Miosic S, Knop K, Holscher D, Greiner J, Gosch C, Thill J, Kai M,

Shrestha BK, Schneider B, Crecelius AC, Schubert US, Svatos

A, Stich K, Halbwirth H (2013) 4-Deoxyaurone formation in

Bidens ferulifolia (Jacq.) DC. PLoS One 8:e61766. doi:10.1371/

journal.pone.0061766

Munoz J, Garcia-Molina F, Varon R, Rodriguez-Lopez JN, Garcia-

Ruiz PA, Garcia-Canovas F, Tudela J (2007) Kinetic charac-

terization of the oxidation of chlorogenic acid by polyphenol

oxidase and peroxidase. Characteristics of the o-quinone. J Agric

Food Chem 55:920–928. doi:10.1021/jf062081

Munoz-Munoz JL, Garcia-Molina F, Garcia-Ruiz PA, Molina-Alar-

con M, Tudela J, Garcia-Canovas F, Rodriguez-Lopez JN (2008)

Phenolic substrates and suicide inactivation of tyrosinase:

kinetics and mechanism. Biochem J 416:431–440. doi:10.1042/

BJ20080892

Munoz-Munoz JL, Garcia-Molina F, Acosta-Motos JR, Arribas E,

Garcia-Ruiz PA, Tudela J, Garcia-Canovas F, Rodriguez-Lopez

JN (2011) Indirect inactivation of tyrosinase in its action on

tyrosine. Acta Biochim Pol 58:477–488

Munoz-Munoz JL, Berna J, Garcia-Molina F, Garcia-Ruiz PA, Tudela

J, Rodriguez-Lopez JN, Garcia-Canovas F (2012a) Unravelling

the suicide inactivation of tyrosinase: a discrimination between

mechanisms. J Mol Catal B Enzym 75:11–19. doi:10.1016/j.

molcatb.2011.11.001

Munoz-Munoz JL, Garcia-Molina F, Berna J, Garcia-Ruiz PA, Varon

R, Tudela J, Rodriguez-Lopez JN, Garcia-Canovas F (2012b)

Kinetic characterisation of o-aminophenols and aromatic o-

diamines as suicide substrates of tyrosinase. Biochim Biophys

Acta 1824:647–655. doi:10.1016/j.bbapap.2012.02.001

Nakayama T (2002) Enzymology of aurone biosynthesis. J Biosci

Bioeng 94:487–491

Nakayama T, Yonekura-Sakakibara K, Sato T, Kikuchi S, Fukui Y,

Fukuchi-Mizutani M, Ueda T, Nakao M, Tanaka Y, Kusumi T,

Nishino T (2000) Aureusidin synthase: a polyphenol oxidase

homolog responsible for flower coloration. Science

290:1163–1166. doi:10.1126/science.290.5494.1163

Nakayama T, Sato T, Fukui Y, Yonekura-Sakakibara K, Hayashi H,

Tanaka Y, Kusumi T, Nishino T (2001) Specificity analysis and

mechanism of aurone synthesis catalyzed by aureusidin syn-

thase, a polyphenol oxidase homolog responsible for flower

coloration. FEBS Lett 499:107–111

Nicholls KW, Bohm BA (1979) Flavonoids and affinities of

Coreopsis bigelovii. Phytochemistry 18:1076. doi:10.1016/

S0031-9422(00)91489-4

536 Planta (2015) 242:519–537

123

Nillius D, Jaenicke E, Decker H (2008) Switch between tyrosinase

and catecholoxidase activity of scorpion hemocyanin by al-

losteric effectors. FEBS Lett 582:749–754. doi:10.1016/j.febslet.

2008.01.056

Okada Y, Okita M, Murai Y, Okano Y, Nomura M (2014) Isolation

and identification of flavonoids from Coreopsis lanceolata L.

petals. Nat Prod Res 28:201–204. doi:10.1080/14786419.2013.

856905

Ono E, Fukuchi-Mizutani M, Nakamura N, Fukui Y, Yonekura-

Sakakibara K, Yamaguchi M, Nakayama T, Tanaka T, Kusumi

T, Tanaka Y (2006a) Yellow flowers generated by expression of

the aurone biosynthetic pathway. Proc Natl Acad Sci USA

103:11075–11080. doi:10.1073/pnas.0604246103

Ono E, Hatayama M, Isono Y, Sato T, Watanabe R, Yonekura-

Sakakibara K, Fukuchi-Mizutani M, Tanaka Y, Kusumi T,

Nishino T, Nakayama T (2006b) Localization of a flavonoid

biosynthetic polyphenol oxidase in vacuoles. Plant J

45:133–143. doi:10.1111/j.1365-313X.2005.02625.x

Ramsden CA, Riley PA (2010a) Mechanistic studies of tyrosinase

suicide inactivation. Arkivoc 1:260–274

Ramsden CA, Riley PA (2010b) Studies of the competing rates of

catechol oxidation and suicide inactivation of tyrosinase.

Arkivoc 10:248–254

Ramsden CA, Riley PA (2014) Tyrosinase: the four oxidation states

of the active site and their relevance to enzymatic activation,

oxidation and inactivation. Bioorg Med Chem 22:2388–2395.

doi:10.1016/j.bmc.2014.02.048

Ramsden CA, Stratford MR, Riley PA (2009) The influence of

catechol structure on the suicide-inactivation of tyrosinase. Org

Biomol Chem 7:3388–3390. doi:10.1039/b910500j

Rollett A, Thallinger B, Ohradanova-Repic A, Machacek C, Walenta

E, Cavaco-Paulo A, Birner-Gruenberger R, Bogner-Strauss JG,

Stockinger H, Guebitz GM (2013) Enzymatic synthesis of

antibody-human serum albumin conjugate for targeted drug

delivery using tyrosinase from Agaricus bisporus. Rsc Advances

3:1460–1467. doi:10.1039/C2ra22560c

Schlangen K, Miosic S, Thill J, Halbwirth H (2010) Cloning,

functional expression, and characterization of a chalcone 3-hy-

droxylase from Cosmos sulphureus. J Exp Bot 61:3451–3459.

doi:10.1093/jxb/erq169

Shakya R, Ye J, Rommens CM (2012) Altered leaf colour is

associated with increased superoxide-scavenging activity in

aureusidin-producing transgenic plants. Plant Biotechnol J

10:1046–1055. doi:10.1111/j.1467-7652.2012.00732.x

Shang YF, Oidovsambuu S, Jeon J-S, Nho CW, Um B-H (2013)

Chalcones from the flowers of Coreopsis lanceolata and their

in vitro antioxidative activity. Planta Med 79:295–300. doi:10.

1055/s-0032-1328188

Shimokoriyama M, Hattori S (1953) Anthochlor pigments of Cosmos

sulphureus, Coreopsis lanceolata and C. saxicola. J Am Chem

Soc 75:1900–1904

Sojo MM, Nunez-Delicado E, Garcia-Carmona F, Sanchez-Ferrer A

(1998) Partial purification of a banana polyphenol oxidase using

triton X-114 and PEG 8000 for removal of polyphenols. J Agric

Food Chem 46:4924–4930. doi:10.1021/Jf980473d

Stover BC, Muller KF (2010) TreeGraph 2: combining and visual-

izing evidence from different phylogenetic analyses. BMC

Bioinformatics 11:7. doi:10.1186/1471-2105-11-7

Strack D, Schliemann W (2001) Bifunctional polyphenol oxidases:

novel functions in plant pigment biosynthesis. Angew Chem Int

Ed Engl 40:3791–3794. doi:10.1002/1521-3773(20011015)40:

20\3791:AID-ANIE3791[3.0.CO;2-T

Tanimoto S, Miyazawa M, Inoue T, Okada Y, Nomura M (2009)

Chemical constituents of Coreopsis lanceolata L. and their

physiological activities. J Oleo Sci 58:141–146. doi:10.5650/jos.

58.141

Tran LT, Constabel CP (2011) The polyphenol oxidase gene family in

poplar: phylogeny, differential expression and identification of a

novel, vacuolar isoform. Planta 234:799–813. doi:10.1007/

s00425-011-1441-9

Tran LT, Taylor JS, Constabel CP (2012) The polyphenol oxidase

gene family in land plants: lineage-specific duplication and

expansion. BMC Genom 13:395. doi:10.1186/1471-2164-13-395

Valero E, Garcia-Carmona F (1992) Hysteresis and cooperative

behavior of a latent plant polyphenoloxidase. Plant Physiol

98:774–776

Valero E, Garcia-Carmona F (1998) pH-Dependent effect of sodium

chloride on latent grape polyphenol oxidase. J Agric Food Chem

46:2447–2451. doi:10.1021/Jf971088b

Virador VM, Reyes Grajeda JP, Blanco-Labra A, Mendiola-Olaya E,

Smith GM, Moreno A, Whitaker JR (2010) Cloning, sequencing,

purification, and crystal structure of Grenache (Vitis vinifera)

polyphenol oxidase. J Agric Food Chem 58:1189–1201. doi:10.

1021/jf902939q

Wahler D, Gronover CS, Richter C, Foucu F, Twyman RM,

Moerschbacher BM, Fischer R, Muth J, Prufer D (2009)

Polyphenoloxidase silencing affects latex coagulation in Tarax-

acum species. Plant Physiol 151:334–346. doi:10.1104/pp.109.

138743

Waite JH (1976) Calculating extinction coefficients for enzymatically

produced o-quinones. Anal Biochem 75:211–218. doi:10.1016/

0003-2697(76)90072-5

Wang W, Chen W, Yang Y, Liu T, Yang H, Xin Z (2014) New

phenolic compounds from Coreopsis tinctoria Nutt. and their

antioxidant and angiotensin I-converting enzyme inhibitory

activities. J Agric Food Chem. doi:10.1021/jf504289g

Yoruk R, Marshall MR (2003) Physicochemical properties and

function of plant polyphenol oxidase: a review1. J Food

Biochem 27:361–422

Zekiri F, Molitor C, Mauracher SG, Michael C, Mayer RL, Gerner C,

Rompel A (2014) Purification and characterization of tyrosinase

from walnut leaves (Juglans regia). Phytochemistry 101:5–15.

doi:10.1016/j.phytochem.2014.02.010

Planta (2015) 242:519–537 537

123